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Modulation of Protein Structure and Function by Lipids
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Modulation of Protein Structure and Function by Lipids

A special issue of International Journal of Molecular Sciences (ISSN 1422-0067). This special issue belongs to the section "Biochemistry".

Deadline for manuscript submissions: closed (31 October 2023) | Viewed by 17170

Special Issue Editor

Dr. Avia Rosenhouse-Dantsker

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Guest Editor
Department of Chemistry, University of Illinois at Chicago, Chicago, IL 60607, USA
Interests: structure–function relationship in proteins; protein–lipid interactions; lipid modulation of protein function; modulation of ion channel function; atrial and neuronal G protein-gated inwardly rectifying potassium channels; cholesterol; phosphoinositides; fatty acids
Special Issues, Collections and Topics in MDPI journals

Special Issue Information

Dear Colleagues,

Since the introduction of the fluid mosaic model of cell membrane structure by Singer and Nicolson in 1972, our understanding of the roles of lipids in cellular functions has evolved significantly. Through insights gained with advances in technology, it has become clear that lipids are not merely passive entities that diffuse freely within membrane bilayers but are also key players in the modulation of protein function. Lipids, such as phosphoinositides, sterols, and fatty acids, have been shown to affect the function of a growing number of proteins (e.g., G protein-coupled receptors, ion channels, transporters, etc.). Lipids exert their effect on proteins through a variety of mechanisms, such as by modulating protein structure, function, and dynamics; promoting protein oligomerization; and mediating protein–protein interactions in the membrane. As a result, lipids have emerged as central players in diverse disease processes and, hence, in drug development, where lipid-regulated proteins are potential drug targets. The aim of this Special Issue is to provide an overview of the current understanding of the impact of lipids on protein structure and function in addition to presenting new developments in this field. Both original papers and reviews on this topic are welcome.

Dr. Avia Rosenhouse-Dantsker
Guest Editor

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Keywords

  • protein modulation by lipids
  • phosphoinositides
  • cholesterol
  • sterols
  • fatty acids
  • G protein-coupled receptors
  • ion channels
  • transporters

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Published Papers (8 papers)

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21 pages, 3559 KiB  
Article
Dimeric Tubulin Modifies Mechanical Properties of Lipid Bilayer, as Probed Using Gramicidin A Channel
by Tatiana K. Rostovtseva, Michael Weinrich, Daniel Jacobs, William M. Rosencrans and Sergey M. Bezrukov
Int. J. Mol. Sci. 2024, 25(4), 2204; https://doi.org/10.3390/ijms25042204 - 12 Feb 2024
Viewed by 902
Abstract
Using the gramicidin A channel as a molecular probe, we show that tubulin binding to planar lipid membranes changes the channel kinetics—seen as an increase in the lifetime of the channel dimer—and thus points towards modification of the membrane’s mechanical properties. The effect [...] Read more.
Using the gramicidin A channel as a molecular probe, we show that tubulin binding to planar lipid membranes changes the channel kinetics—seen as an increase in the lifetime of the channel dimer—and thus points towards modification of the membrane’s mechanical properties. The effect is more pronounced in the presence of non-lamellar lipids in the lipid mixture used for membrane formation. To interpret these findings, we propose that tubulin binding redistributes the lateral pressure of lipid packing along the membrane depth, making it closer to the profile expected for lamellar lipids. This redistribution happens because tubulin perturbs the lipid headgroup spacing to reach the membrane’s hydrophobic core via its amphiphilic α-helical domain. Specifically, it increases the forces of repulsion between the lipid headgroups and reduces such forces in the hydrophobic region. We suggest that the effect is reciprocal, meaning that alterations in lipid bilayer mechanics caused by membrane remodeling during cell proliferation in disease and development may also modulate tubulin membrane binding, thus exerting regulatory functions. One of those functions includes the regulation of protein–protein interactions at the membrane surface, as exemplified by VDAC complexation with tubulin. Full article
(This article belongs to the Special Issue Modulation of Protein Structure and Function by Lipids)
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Figure 1

Figure 1
<p>Tubulin increases the lifetime of grA channels and decreases their conductance in DOPE membranes but not in DOPC membranes: (<b>A</b>) Current traces of grA channels in DOPE membrane before (trace a) and after (trace b) addition of 30 nM tubulin. Tubulin notably increases grA lifetime and decreases channel conductance in the DOPE membrane. Tubulin also induces fast current flickering that can be better seen at a finer time scales in inset d in comparison with the control trace in inset c. (<b>B</b>) Current traces of grA channels in DOPC membrane before (trace a) and after (trace b) addition of 50 nM tubulin. The addition of 50 nM of tubulin does not appreciably change grA channel parameters in the DOPC membranes. The applied voltage was 100 mV. Tubulin was added to the <span class="html-italic">cis</span> compartment. Current records were digitally filtered using an averaging time of 10 ms. Dashed lines indicate zero current level and dotted lines indicate the currents through single grA channels. Here (and elsewhere) the medium consisted of 1 M KCl buffered with 5 mM HEPES at pH 7.4.</p> Full article ">Figure 2
<p>Effect of tubulin on the lifetime and conductance of grA channels, which depends on DOPE content in DOPE/DOPC mixture. In DOPE membranes, tubulin increases grA lifetime (<b>A</b>) and decreases conductance (<b>B</b>) in a dose-dependent manner that displays saturation at about 20 nM tubulin concentration. There is virtually no effect of tubulin on the channel lifetime and conductance in DOPC membranes. Channel conductance is given as its ratio in the presence of tubulin to that in the absence of tubulin. (<b>C</b>,<b>D</b>) Effect of 30 nM tubulin on the channel lifetime (<b>C</b>) and conductance (<b>D</b>) increases with PE content in the PE/PC mixture. Lines are drawn to guide the eye. Data are the mean values obtained in 3–5 experiments ± S.E. Experimental conditions are as in <a href="#ijms-25-02204-f001" class="html-fig">Figure 1</a>.</p> Full article ">Figure 3
<p>Tubulin induces fast flickering of the ionic current through grA channels in a dose- and voltage-dependent manner: (<b>A</b>) grA current traces were obtained at indicated tubulin concentrations in the DPhPC membrane at 200 mV applied voltage. The flickering increases with tubulin concentration and the corresponding decrease of the average conductance (black dotted line indicates conductance at 0 and 3 nM tubulin and red dotted line indicates conductance at 10 and 50 nM tubulin). Current records were filtered with a digital 8-pole Bessel filter at 1 kHz. (<b>B</b>) Power spectral densities of current fluctuations at 50 nM of tubulin (upper trace) can be approximated by a Lorentzian spectrum (smooth line through the data) with a corner frequency of <span class="html-italic">f<sub>c</sub></span> = 600 Hz. (<b>C</b>) Tubulin-induced current fluctuations in the grA channel increase with applied voltage. Power spectral densities of current fluctuations of the single grA channel obtained at applied voltages as indicated in the presence of 50 nM tubulin. Solid lines are fitted with Lorentzian spectra. Current records were filtered with a digital 8-pole Bessel filter at 2 kHz. The medium consisted of 1 M KCl buffered with 5 mM HEPES at pH 7.4.</p> Full article ">Figure 4
<p>Tubulin-S changes the lifetime and conductance of grA channels in DOPE membranes. Similarly to tubulin, tubulin-S increases channel lifetime (<b>A</b>) and reduces its conductance (<b>B</b>) in comparison with the control. Channel lifetime and relative conductance (<span class="html-italic">G<sub>Tub</sub></span>/<span class="html-italic">G<sub>Cntr</sub></span>) were measured in the presence of 50 nM tubulin and 40 nM tubulin-S in the <span class="html-italic">cis</span> compartment. Data are the mean values obtained in 3–4 experiments ± S.E. Other experimental conditions were as in <a href="#ijms-25-02204-f001" class="html-fig">Figure 1</a>.</p> Full article ">Figure 5
<p>Tubulin-induced fast flickering of grA channels in diC(22:1)PC bilayers. (<b>A</b>) Current traces of a single grA channel in a diC(22:1)PC bilayer before (trace a) and after (trace b) addition of 30 nM tubulin to the <span class="html-italic">cis</span> compartment. The addition of tubulin induces rapid events of grA channel closure to a zero-current level (indicated by black dashed line), as shown in trace c at a finer time scale. The dotted black line indicates open channel conductance. The applied voltage was 200 mV. Current records were filtered with a digital 8-pole Bessel filter at 2 kHz. (<b>B</b>) Power spectral density of tubulin-induced current fluctuations, which depends on the polarity of the applied voltage. Solid gray lines represent the fits to Lorentzian spectra.</p> Full article ">Figure 6
<p>Illustration of stable α-tubulin binding to the DOPE membrane surfaces from ~800 ns of all-atom ANTON MD simulations. The insertion helical region (A330–W346) is colored according to chemical functionality (negative residues in red, positive residues in blue, and hydrophobic/aromatic in orange). The location of W346 is shown in sphere mode. β-tubulin, shown in the magenta-colored ribbon, is added based on RMSF alignment for α-tubulin in the dimer. MD simulations were performed for α-tubulin only. The unstructured C-terminal tails are not shown. Adapted with permission from Hoogerheide et al., <span class="html-italic">Proc. Natl. Acad. Sci. USA</span> (2017) [<a href="#B29-ijms-25-02204" class="html-bibr">29</a>]. Copyright (2017) National Academy of Sciences.</p> Full article ">Figure 7
<p>α-Tubulin peptide increases grA lifetime: (<b>A</b>) Current traces of grA channels in DOPE membrane before (Control) and after the addition of 10 and 20 μM of α-tubulin peptide to both sides of the DOPE membrane. Current records were filtered with a digital 8-pole Bessel filter at 1 kHz. Dashed lines indicate zero current level. The membrane bathing solution contained 1 M KCl buffered with 5 mM HEPES at pH 7.4. The applied voltage was 100 mV. α-tubulin peptide was dissolved in DMSO. (<b>B</b>) α-tubulin peptide increases grA lifetime in a dose-dependent manner. Bars and error bars are the mean and standard deviation from the mean; the symbols represent data points of 4 independent experiments. Control measurements with the addition of DMSO aliquots corresponding to 20 μM of α-tubulin peptide addition do not show the effect on grA lifetime.</p> Full article ">Figure 8
<p>Binding curves for α-tubulin peptide interaction with DOPE and DOPC membranes, as measured by BOA. The transmembrane potential ΔΨ changes due to α-tubulin peptide binding to the DOPE/DOPC (4:1) (reddish symbols) membranes and does not change for DOPC (bluish symbols) membranes. Membranes were formed in 150 mM KCl buffered with 5 mM HEPES at pH 7.4. α-Tubulin peptide dissolved in DMSO was added to the <span class="html-italic">cis</span> side of the membrane. Aliquots of DMSO were correspondingly added to the <span class="html-italic">trans</span> side. Large red and blue circles and error bars are the mean and standard deviation from the mean for DOPE/DOPC and DOPC membranes, respectively; they represent data points of 7 individual experiments for DOPE/DOPC membranes and 6 experiments for DOPC membranes. The solid line is a fit to the binding equation with <span class="html-italic">K<sub>d</sub></span> = 156 μM.</p> Full article ">Figure 9
<p>Schematics of the effect of tubulin dimer on grA lifetime and conductance: (<b>A</b>) Binding of α-β-tubulin heterodimers to the DOPE membrane reduces packing stress of the lipid tails, which is observed as the increase of grA channel (shown in red) lifetime. (<b>B</b>) In the case of diC(22:1)PC membranes, binding of the tubulin dimers is limited to the regions of membranes where headgroup packing is distorted by grA channel presence in the region of the lipid funnel forming the entrance to the channel. This limitation leads to the unchanged integral properties of the membrane and unchanged grA lifetime (<a href="#ijms-25-02204-t001" class="html-table">Table 1</a>); however, the localized binding is clearly manifested via transient channel blockages (<a href="#ijms-25-02204-f005" class="html-fig">Figure 5</a>) from the bulky body of the tubulin dimer. Created with <a href="http://Biorender.com" target="_blank">Biorender.com</a>.</p> Full article ">
24 pages, 15209 KiB  
Article
Cross-Linking Mass Spectrometry on P-Glycoprotein
by Gabriella Gellen, Eva Klement, Kipchumba Biwott, Gitta Schlosser, Gergő Kalló, Éva Csősz, Katalin F. Medzihradszky and Zsolt Bacso
Int. J. Mol. Sci. 2023, 24(13), 10627; https://doi.org/10.3390/ijms241310627 - 25 Jun 2023
Cited by 1 | Viewed by 2322
Abstract
The ABC transporter P-glycoprotein (Pgp) has been found to be involved in multidrug resistance in tumor cells. Lipids and cholesterol have a pivotal role in Pgp’s conformations; however, it is often difficult to investigate it with conventional structural biology techniques. Here, we applied [...] Read more.
The ABC transporter P-glycoprotein (Pgp) has been found to be involved in multidrug resistance in tumor cells. Lipids and cholesterol have a pivotal role in Pgp’s conformations; however, it is often difficult to investigate it with conventional structural biology techniques. Here, we applied robust approaches coupled with cross-linking mass spectrometry (XL-MS), where the natural lipid environment remains quasi-intact. Two experimental approaches were carried out using different cross-linkers (i) on living cells, followed by membrane preparation and immunoprecipitation enrichment of Pgp, and (ii) on-bead, subsequent to membrane preparation and immunoprecipitation. Pgp-containing complexes were enriched employing extracellular monoclonal anti-Pgp antibodies on magnetic beads, followed by on-bead enzymatic digestion. The LC-MS/MS results revealed mono-links on Pgp’s solvent-accessible residues, while intraprotein cross-links confirmed a complex interplay between extracellular, transmembrane, and intracellular segments of the protein, of which several have been reported to be connected to cholesterol. Harnessing the MS results and those of molecular docking, we suggest an epitope for the 15D3 cholesterol-dependent mouse monoclonal antibody. Additionally, enriched neighbors of Pgp prove the strong connection of Pgp to the cytoskeleton and other cholesterol-regulated proteins. These findings suggest that XL-MS may be utilized for protein structure and network analyses in such convoluted systems as membrane proteins. Full article
(This article belongs to the Special Issue Modulation of Protein Structure and Function by Lipids)
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Graphical abstract

Graphical abstract
Full article ">Figure 1
<p>Regions of Pgp on its primary and tertiary structures. (<b>A</b>) Primary structure of Pgp visualized using Protter 1.0 [<a href="#B22-ijms-24-10627" class="html-bibr">22</a>]. Regions of the protein indicated in pink form nucleotide-binding site 1 (NBS1) and the ones indicated in brown belong to NBS2. TMH 1–12 are indicated in blue. Cholesterols identified in the 6qex PDB structure are labeled in yellow. (<b>B</b>) Tertiary structure of Pgp using inward-facing human Pgp structure, 6qex, with its specific regions indicated. TMD1 (light blue) connected to NBD1 (dark blue), TMD2 (pale green) connected to NBD2 (dark green), NBS1 regions (pink), NBS2 (brown) and flexible linker and elbow helices (grey). N-glycans on ECL1 are blue, while oxygen atoms are labeled in red (see abbreviations in the text of the Introduction).</p> Full article ">Figure 1 Cont.
<p>Regions of Pgp on its primary and tertiary structures. (<b>A</b>) Primary structure of Pgp visualized using Protter 1.0 [<a href="#B22-ijms-24-10627" class="html-bibr">22</a>]. Regions of the protein indicated in pink form nucleotide-binding site 1 (NBS1) and the ones indicated in brown belong to NBS2. TMH 1–12 are indicated in blue. Cholesterols identified in the 6qex PDB structure are labeled in yellow. (<b>B</b>) Tertiary structure of Pgp using inward-facing human Pgp structure, 6qex, with its specific regions indicated. TMD1 (light blue) connected to NBD1 (dark blue), TMD2 (pale green) connected to NBD2 (dark green), NBS1 regions (pink), NBS2 (brown) and flexible linker and elbow helices (grey). N-glycans on ECL1 are blue, while oxygen atoms are labeled in red (see abbreviations in the text of the Introduction).</p> Full article ">Figure 2
<p>Experimental approaches for XL-MS on Pgp. Cross-linking with lysine-reactive cross-linkers (DSSO and BS2Gd<sub>0</sub>/d<sub>4</sub>) was carried out either on living cells followed by membrane preparation, or on-bead after membrane preparation, then affinity purification. Cross-linked complexes containing Pgp were enriched via extracellular mAbs (15D3 and UIC2). Peptide generation via trypsin digestion was performed on-bead, and measurement of samples was realized via LC-MS/MS. Steps involved only in the living cell approach are indicated in pink, and the on-bead experimental steps are brown, while steps applied in both approaches are indicated in black.</p> Full article ">Figure 3
<p>The binding of the (<b>A</b>) 15D3 and (<b>B</b>) UIC-2 mAbs increased after PNGase F treatment applied at different concentrations on living cells as measured via flow cytometry. N-glycosyl groups sterically affect the docking of both antibodies to their binding site. (<b>C</b>) Western blot analysis indicates a small shift in the electrophoretic mobility of the Pgp protein after deglycosylation.</p> Full article ">Figure 4
<p>Overlap of unique mono-links on P-glycoprotein in different cross-linking setups. Each type of sample preparation was performed employing UIC2 and 15D3 monoclonal antibodies. Out of 34 unique mono-links, 6 (17.6%) were found in all sample preparation approaches; these are the ones that are most readily solvent accessible. In total, 16 (8 + 6 + 2, 47%) of the mono-links were identified at least with two different sample preparation modes which suggests that, even with distinct methods, it is possible to identify a similar set of modifications. Mono-links detected via each experimental approach visualized on Pgp’s primary structure using Protter 1.0 and on Pgp’s tertiary 6qex PDB structure can be found in <a href="#app1-ijms-24-10627" class="html-app">Figures S1 and S2</a> accordingly.</p> Full article ">Figure 5
<p>Solvent accessibility of lysines on Pgp. (<b>A</b>) Electrostatic potential surface representation of inward-facing Pgp (6qex); nitrogen atoms of all lysines are represented as grey balls, and buried ones are colored in yellow. Five lysines with a legend are the ones which were modified by cross-linkers. (<b>B</b>) Hydrophobicity and charge distribution on Pgp’s surface. Carbon atoms not bound to nitrogen or oxygen atoms are yellow, oxygens with negative charges in glutamate and aspartate are red and nitrogens carrying positive charges in lysine and arginine are blue, while all other atoms are white [<a href="#B51-ijms-24-10627" class="html-bibr">51</a>]. The inlet shows extracellular lysines of which only K734 and K967 are surface exposed; however, when Pgp is in complex with UIC2 or 15D3 they are also buried.</p> Full article ">Figure 6
<p>Mono-links and cross-links on Pgp. (<b>A</b>) Overlap of cross-links between different sample preparation approaches. Eleven cross-links were identified with all sample preparation approaches, with a 27% overlap between the living cell and the on-bead DSSO methods. No cross-links were identified with the living cell BS2Gd<sub>0</sub>/d<sub>4</sub> approach. (<b>B</b>) Linear plots of cross-linked sites on human Pgp identified with on-bead and living cell approaches. All lysines are indicated in light blue, and cross-links are purple. (<b>C</b>) Mono-links highlighted in green balls and cross-links indicated by red lines on the structure of Pgp in complex with UIC2 antibody (6qex) based on the results of the cross-linking approaches. Lysines which are not mono-linked are depicted as grey balls, and cholesterols and lipids are yellow. The inlet shows the ICH3 loop involved in cross-links highlighted in pink, which was previously described to be influenced by the presence of cholesterol. (<b>D</b>) Mono-links of Walker A regions and other segments of nucleotide binding sites (NBS) emphasized in pink. (<b>E</b>) ICH3 directly connected to ECL5, cross-linked to C-loop, which has cross-link with ICH4, directly connected to ECL6. Connections of these regions on Pgp suggest a complex interplay between segments that have previously been described as cholesterol-sensitive. The inlet highlights the cholesterol-sensitive binding sites of 15D3 mAb, and mono-linked lysines are colored in green. One mono-link on 15D3 fell in the binding region close to Pgp’s K967 lysine on ECL6; however, cross-link formation between them could not be detected unambiguously.</p> Full article ">Figure 7
<p>Pgp K786 is cross-linked to K826. (<b>A</b>) The MS2-CID spectrum dominated by the diagnostic peptide pairs of the two cross-linked peptides formed upon fragmentation along the cross-linker. (<b>B</b>) The MS2-HCD spectrum containing additional fragments of the two peptides along the peptide backbone to assist identification. Fragments labeled red and orange belong to Peptide A while fragments labeled blue and light blue belong to Peptide B. (<b>C</b>–<b>F</b>) MS3-HCD spectra of the peptide pairs detected in MS2-CID confirm the identity of Peptide A (<b>C</b>,<b>D</b>) and Peptide B (<b>E</b>,<b>F</b>). Peptide A is LANDAAQV<b>K</b>GAIGSR [818–832], and Peptide B is AGEILT<b>K</b>R [780–787]. All other cross-link spectra are available in the <a href="#app1-ijms-24-10627" class="html-app">Supplementary Materials (Figures S5–S15)</a>.</p> Full article ">Figure 8
<p>ClusPro 2.0 prediction of docking (<b>A</b>) 15D3 mAb to Pgp (6qex); the inlet indicates binding to ECL1, ECL5 and ECL6, partially overlapping with the epitopes of (<b>B</b>) UIC2 (ECL1, ECL3, and ECL4), visualized by means of the UIC2-associated, 6qex PDB structure. ECLs of Pgp are highlighted in pink. NHS cross-linkers could potentially target K748 on ECL4 and K967 on ECL6; however, we did not see any modifications in these lysine residues, probably due to the vicinity of the membrane and the antibodies embedding the lysine side chains.</p> Full article ">Figure 9
<p>Differences between 15D3 and UIC2 IP enrichments. (<b>A</b>) Overlap of protein identifications between the two mAbs. (<b>B</b>,<b>C</b>) Protein interaction networks of Pgp enriched by 15D3 (<b>B</b>) and UIC2 (<b>C</b>) mAbs. Interactions were gathered and depicted using the STRING database employing all interaction sources [<a href="#B71-ijms-24-10627" class="html-bibr">71</a>] and Cytoscape software accordingly. Only proteins with at least 10 SPCs were considered. Pgp is labeled in yellow, its well-known protein partners and proteins related to MDR are highlighted in red and neighbors of these highlighted nodes are pale grey. The nodes’ size is proportional to the individual proteins’ SPCs. SPCs were normalized to all peptide-spectrum matches (PSMs) in the given analysis. Nodes of the large and small ribosome subunits and components of the chaperonin-containing T-complex were grouped together to reduce the complexity of the network. Pgp was approximately 7 times more abundant based on SPCs using UIC2 for the IP and pulled down 222 proteins, whereas with 15D3 IP a more diverse set of 368 proteins could be enriched, many of which were cholesterol-sensitive. These networks indicate that enriched proteins are indeed related to Pgp. (<b>D</b>–<b>F</b>) Cellular component distribution of 15D3 enrichment (<b>D</b>), the 160 proteins uniquely identified in the 15D3 pull-down experiment (<b>E</b>), and proteins enriched in the UIC2 immunoprecipitation experiment (<b>F</b>) analyzed with the ClueGO app within Cytoscape [<a href="#B72-ijms-24-10627" class="html-bibr">72</a>].</p> Full article ">
16 pages, 1671 KiB  
Article
Differential Functional Contribution of BK Channel Subunits to Aldosterone-Induced Channel Activation in Vascular Smooth Muscle and Eventual Cerebral Artery Dilation
by Steven C. Mysiewicz, Sydney M. Hawks, Anna N. Bukiya and Alex M. Dopico
Int. J. Mol. Sci. 2023, 24(10), 8704; https://doi.org/10.3390/ijms24108704 - 12 May 2023
Viewed by 1577
Abstract
Calcium/voltage-activated potassium channels (BK) control smooth muscle (SM) tone and cerebral artery diameter. They include channel-forming α and regulatory β1 subunits, the latter being highly expressed in SM. Both subunits participate in steroid-induced modification of BK activity: β1 provides recognition for [...] Read more.
Calcium/voltage-activated potassium channels (BK) control smooth muscle (SM) tone and cerebral artery diameter. They include channel-forming α and regulatory β1 subunits, the latter being highly expressed in SM. Both subunits participate in steroid-induced modification of BK activity: β1 provides recognition for estradiol and cholanes, resulting in BK potentiation, whereas α suffices for BK inhibition by cholesterol or pregnenolone. Aldosterone can modify cerebral artery function independently of its effects outside the brain, yet BK involvement in aldosterone’s cerebrovascular action and identification of channel subunits, possibly involved in steroid action, remains uninvestigated. Using microscale thermophoresis, we demonstrated that each subunit type presents two recognition sites for aldosterone: at 0.3 and ≥10 µM for α and at 0.3–1 µM and ≥100 µM for β1. Next, we probed aldosterone on SM BK activity and diameter of middle cerebral artery (MCA) isolated from β1−/− vs. wt mice. Data showed that β1 leftward-shifted aldosterone-induced BK activation, rendering EC50~3 μM and ECMAX ≥ 10 μM, at which BK activity increased by 20%. At similar concentrations, aldosterone mildly yet significantly dilated MCA independently of circulating and endothelial factors. Lastly, aldosterone-induced MCA dilation was lost in β1−/− mice. Therefore, β1 enables BK activation and MCA dilation by low µM aldosterone. Full article
(This article belongs to the Special Issue Modulation of Protein Structure and Function by Lipids)
►▼ Show Figures

Figure 1

Figure 1
<p>(<b>A</b>) Sketch of a BK channel heterodimer of α (slo1) (orange) and generic β (types 1–4) (green) subunits, with pore in the former between S5 and S6. Four heterodimers make the functional BK channels expressed in most mammalian tissues. The effect of different steroids on BK channel activity is shown, with the number of carbons in the steroid molecule and the BK subunit involved in steroid action in parenthesis. EC: extracellular medium; IC: intracellular medium S: transmembrane segment in α; RCK: regulator of conductance for K<sup>+</sup> domain; TM: transmembrane domain in β; DHEA: dehydroepiandrosterone. (<b>B</b>) Structure of aldosterone (C21).</p> Full article ">Figure 2
<p>Differential scanning fluorometry on isolated BK-associated proteins shows differential binding of aldosterone. (<b>A</b>) Example of normalized traces from a single thermal unfolding run of isolated BK α subunit (slo1; cbv1 isoform). In each run, protein samples were loaded in parallel into multiple capillaries with various aldosterone concentrations. (<b>B</b>) Differences between onset and inflection point of averaged cbv1 curves show significant differences in unfolding at 0.3, 10, and 100 µM aldosterone. Average values reflect data from no fewer than three runs representing biological replicates. (<b>C</b>) Example of normalized traces from a single thermal unfolding run of isolated BK β<sub>1</sub> subunit. As in (<b>B</b>), during one run, protein samples were loaded in parallel into multiple capillaries with various aldosterone concentrations. (<b>D</b>) Differences between onset and inflection point of averaged β<sub>1</sub> curves show a significant difference at 0.3 and at 100 µM aldosterone. * <span class="html-italic">p</span> &lt; 0.05 unless otherwise stated. Average values reflect data from no fewer than three runs representing biological replicates.</p> Full article ">Figure 3
<p>Aldosterone activates smooth muscle BK channels from <span class="html-italic">wt</span> mouse middle cerebral artery (MCA) in a concentration-dependent manner; this action is blunted in myocyte BK channels from <span class="html-italic">KCNMB1<sup>−/−</sup></span> mice. (<b>A</b>) Representative channel activity traces before (top), during (middle), and after (bottom) application of 3 µM aldosterone to the cytosolic side of inside-out (I/O) patches from <span class="html-italic">KCNMB1<sup>−/−</sup></span> myocytes. (<b>B</b>) In these myocytes, channel steady-state activity (NPo) as a function of aldosterone concentration shows a mild increase only at 100 µM. (<b>C</b>) Channel activity traces before (top), during (middle), and after (bottom) application of 3 µM aldosterone to the cytosolic side of I/O patches from <span class="html-italic">wt</span> myocytes. (<b>D</b>) In <span class="html-italic">wt</span> cells, aldosterone action is concentration-dependent: EC<sub>50</sub> = 3 µM; EC<sub>MAX</sub> = 10 µM. In (<b>A</b>,<b>C</b>): red dotted lines and an oblique red arrow indicate the baseline (all channels closed).</p> Full article ">Figure 4
<p>Aldosterone mildly dilates in vitro pressurized, de-endothelialized middle cerebral arteries (MCA) from <span class="html-italic">wt</span> mice but fails to do so in MCA from <span class="html-italic">KCNMB1<sup>−/−</sup></span> mice. (<b>A</b>,<b>B</b>) Representative MCA diameter traces from <span class="html-italic">KCNMB1<sup>−/−</sup></span> mouse showing diameter changes in response to aldosterone or depolarizing 60 mM KCl. (<b>C</b>,<b>D</b>) Corresponding results from <span class="html-italic">wt</span> mice show that aldosterone mildly dilates MCA in a concentration-dependent manner.</p> Full article ">Figure 5
<p>In intact (+endothelium) middle cerebral arteries (MCA) from <span class="html-italic">wt</span> mice, aldosterone-induced dilation is reduced when compared to that in de-endothelialized MCA (<a href="#ijms-24-08704-f004" class="html-fig">Figure 4</a>). (<b>A</b>,<b>B</b>) Representative MCA diameter traces from <span class="html-italic">KCNMB1<sup>−/−</sup></span> mouse showing diameter changes in response to aldosterone or depolarizing 60 mM KCl. (<b>C</b>,<b>D</b>) Corresponding results from <span class="html-italic">wt</span> mice show a very small dilation by &gt;0.3 μM aldosterone.</p> Full article ">
21 pages, 4213 KiB  
Article
Heart-Type Fatty Acid Binding Protein Binds Long-Chain Acylcarnitines and Protects against Lipotoxicity
by Diana Zelencova-Gopejenko, Melita Videja, Aiga Grandane, Linda Pudnika-Okinčica, Anda Sipola, Karlis Vilks, Maija Dambrova, Kristaps Jaudzems and Edgars Liepinsh
Int. J. Mol. Sci. 2023, 24(6), 5528; https://doi.org/10.3390/ijms24065528 - 14 Mar 2023
Cited by 1 | Viewed by 1962
Abstract
Heart-type fatty-acid binding protein (FABP3) is an essential cytosolic lipid transport protein found in cardiomyocytes. FABP3 binds fatty acids (FAs) reversibly and with high affinity. Acylcarnitines (ACs) are an esterified form of FAs that play an important role in cellular energy metabolism. However, [...] Read more.
Heart-type fatty-acid binding protein (FABP3) is an essential cytosolic lipid transport protein found in cardiomyocytes. FABP3 binds fatty acids (FAs) reversibly and with high affinity. Acylcarnitines (ACs) are an esterified form of FAs that play an important role in cellular energy metabolism. However, an increased concentration of ACs can exert detrimental effects on cardiac mitochondria and lead to severe cardiac damage. In the present study, we evaluated the ability of FABP3 to bind long-chain ACs (LCACs) and protect cells from their harmful effects. We characterized the novel binding mechanism between FABP3 and LCACs by a cytotoxicity assay, nuclear magnetic resonance, and isothermal titration calorimetry. Our data demonstrate that FABP3 is capable of binding both FAs and LCACs as well as decreasing the cytotoxicity of LCACs. Our findings reveal that LCACs and FAs compete for the binding site of FABP3. Thus, the protective mechanism of FABP3 is found to be concentration dependent. Full article
(This article belongs to the Special Issue Modulation of Protein Structure and Function by Lipids)
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Figure 1

Figure 1
<p>(<b>A</b>) Structure of heart-type fatty acid-binding protein (FABP3, PDB ID 3WVM [<a href="#B15-ijms-24-05528" class="html-bibr">15</a>]) in complex with palmitate (C16:0). Protein structure is represented as ribbons and colored according to the secondary structure elements as follows: α-helixes shown in orange, β-sheets in cyan, and loops in gray. C16:0 is shown as dark blue sticks, while residues involved in ligand and water cluster binding are shown as light gray balls and sticks. Water molecules (red spheres) and hydrogen bonds (yellow dashed lines) are shown. (<b>B</b>) Human FABP3 amino acid sequence. Residues colored in yellow were not assigned from the NMR spectra either for the apo-form or for the holo-form, while residues in cyan were not assigned only for complexes of FABP3 with acylcarnitines (ACs). Crosspeaks for residues colored in gray were not observed in any 2D <sup>1</sup>H-<sup>15</sup>N HSQC spectra. The black arrow points to the position of the TEV cleavage site. (<b>C</b>) Assigned 2D <sup>1</sup>H-<sup>15</sup>N HSQC spectra of human apo-FABP3 in 20 mM K<sub>2</sub>HPO<sub>4</sub>/KH<sub>2</sub>PO<sub>4</sub>, 50 mM KCl buffer pH 7.6 (KPi). His-tagged FABP3 (noncleaved) is shown in blue, and cleaved FABP3 is shown in red. Backbone amide resonances are denoted as one letter symbol and residue number according to the sequence in <a href="#ijms-24-05528-f001" class="html-fig">Figure 1</a>B. Labels for the residues from the His-tag are shown in light blue. Side chain amide resonances were not assigned. Residues are numbered according to UniProt ID P05413.</p> Full article ">Figure 2
<p>Toxicity of C16:0-carnitine in PANC-1 cells after 4 h of incubation in the presence or in the absence of 60 μM heart-type fatty acid-binding protein (FABP3) in the cell media. Data are shown as the mean ± SEM of 3 independent experiments in at least 6 technical replicates. * indicates a significant difference compared to the cells not subjected to FABP3 treatment at the respective C16:0-carnitine concentration.</p> Full article ">Figure 3
<p>Toxicity of C16:0-carnitine in native and heart-type fatty acid-binding protein (FABP3)-overexpressing PANC-1 cells after 4 h of incubation. Data are shown as the mean ± SEM of 3 independent experiments in at least 6 technical replicates. * indicates a significant difference compared to the native cells at the respective concentration of C16:0-carnitine.</p> Full article ">Figure 4
<p>Superposition of the 2D <sup>1</sup>H-<sup>15</sup>N HSQC spectra for (<b>A</b>) heart-type fatty acid-binding protein (FABP3)-C18:1(n-9)<span class="html-italic">t</span> and (<b>B</b>) FABP3-C18:1(n-9)<span class="html-italic">t</span>-carnitine complex in 20 mM K<sub>2</sub>HPO<sub>4</sub>/KH<sub>2</sub>PO<sub>4</sub>, 50 mM KCl buffer pH 7.6 (KPi). The spectrum of apo-FABP3—in blue, spectrum of the FABP3-ligand complex—in red. Residues with chemical shift perturbations (CSPs) larger than the mean plus one standard deviation are assigned and black arrows show the shift of the corresponding crosspeak. * marks the crosspeaks that have disappeared upon binding of the acylcarnitines (ACs).</p> Full article ">Figure 5
<p>Mapping of the chemical shift perturbations (CSPs) caused by binding of (<b>A</b>) C8:0, (<b>B</b>) C12:0, (<b>C</b>) C14:0, (<b>D</b>) C16:0, (<b>E</b>) C18:1(n-9)<span class="html-italic">c</span>, (<b>F</b>) C20:5(n-3)<span class="html-italic">c</span>, (<b>G</b>) C8:0-carnitine, (<b>H</b>) C12:0-carnitine, (<b>I</b>) C14:0-carnitine, (<b>J</b>) C16:0-carnitine, (<b>K</b>) C18:1(n-9)<span class="html-italic">c</span>-carnitine, or (<b>L</b>) C20:5(n-3)<span class="html-italic">c</span>-carnitine onto the heart-type fatty acid-binding protein (FABP3) structure. The CSPs are color-coded, in which red indicates larger shifts, while blue indicates no changes in the averaged <span class="html-italic">δ<sub>H</sub></span> and <span class="html-italic">δ<sub>N</sub></span> NMR chemical shifts. Unassigned or disappeared residues are colored in gray. FABP3 structure is taken from PDB ID 3WVM.</p> Full article ">Figure 6
<p>(<b>A</b>) Superposition of the ITC titration curves of the heart-type fatty acid-binding protein (FABP3) interaction with C18:1(n-9)<span class="html-italic">c</span> in black and C18:1(n-9)<span class="html-italic">c</span>-carnitine in cyan. Both experiments were performed in 20 mM K<sub>2</sub>HPO<sub>4</sub>/KH<sub>2</sub>PO<sub>4</sub>, 50 mM KCl buffer pH 7.6 (KPi) at 25 °C. Graphical representation of the thermodynamic binding parameters of the FABP3 interaction with (<b>B</b>) C18:1(n-9)<span class="html-italic">c</span> and (<b>C</b>) C18:1(n-9)<span class="html-italic">c</span>-carnitine.</p> Full article ">Figure 7
<p>Superposition of the ITC thermograms for heart-type fatty acid-binding protein (FABP3) titrated with (<b>A</b>) C18:1(n-9)<span class="html-italic">c</span> and (<b>B</b>) C18:1(n-9)<span class="html-italic">c</span>-carnitine at three different temperatures: 16 (cyan), 25 (black), and 37 °C (red).</p> Full article ">Figure 8
<p>Heat capacity change, Δ<span class="html-italic">C<sub>p</sub></span>, and temperature dependence of the binding free energy, Δ<span class="html-italic">G</span>. (<b>A</b>) Δ<span class="html-italic">C<sub>p</sub></span> values for four fatty acids (FAs) and corresponding acylcarnitines (ACs) binding to heart-type fatty acid-binding protein (FABP3). (<b>B</b>) Δ<span class="html-italic">G</span> of FA or AC binding to FABP3 within the temperature range from 0 to 100 °C.</p> Full article ">Figure 9
<p>Top. Competitive binding studies of heart-type fatty acid-binding protein (FABP3)-C18:1(n-9)<span class="html-italic">t</span>-carnitine and three fatty acids (FAs) of different chain lengths. (<b>A</b>) Reference titration of C18:1(n-9)<span class="html-italic">t</span>-carnitine to apo-FABP3. (<b>B</b>) FABP3-C8:0 complex titrated with C18:1(n-9)<span class="html-italic">t</span>-carnitine. (<b>C</b>) FABP3-C10:0 complex titrated with C18:1(n-9)<span class="html-italic">t</span>-carnitine. (<b>D</b>) FABP3-C18:1(n-9)<span class="html-italic">t</span>-carnitine complex titrated with C10:0. (<b>E</b>) FABP3-C18:1(n-9)<span class="html-italic">t</span>-carnitine complex titrated with C12:0. All experiments were performed in 20 mM K<sub>2</sub>HPO<sub>4</sub>/KH<sub>2</sub>PO<sub>4</sub> and 50 mM KCl buffer pH 7.6 (KPi) at 25 °C. Bottom. Schematic representation of ligand binding and competition. Protein is represented as the dark blue sector, acylcarnitines (ACs) as orange, and FAs as cyan circles. Green arrows indicate that the competition event between ligands was successful.</p> Full article ">Scheme 1
<p>General scheme of the synthesis of saturated <b>12</b>–<b>15</b> and unsaturated <b>16</b>–<b>18</b> acylcarnitines (ACs).</p> Full article ">
21 pages, 3296 KiB  
Article
Role of Thylakoid Lipids in Protochlorophyllide Oxidoreductase Activation: Allosteric Mechanism Elucidated by a Computational Study
by Ruiyuan Liu, Leng Wang, Yue Meng, Fang Li, Haiyu Nie and Huizhe Lu
Int. J. Mol. Sci. 2023, 24(1), 307; https://doi.org/10.3390/ijms24010307 - 24 Dec 2022
Cited by 1 | Viewed by 1575
Abstract
Light-dependent protochlorophyllide oxidoreductase (LPOR) is a chlorophyll synthetase that catalyzes the reduction of protochlorophyllide (Pchlide) to chlorophyllide (Chlide) with indispensable roles in regulating photosynthesis processes. A recent study confirmed that thylakoid lipids (TL) were able to allosterically enhance modulator-induced LPOR activation. However, the [...] Read more.
Light-dependent protochlorophyllide oxidoreductase (LPOR) is a chlorophyll synthetase that catalyzes the reduction of protochlorophyllide (Pchlide) to chlorophyllide (Chlide) with indispensable roles in regulating photosynthesis processes. A recent study confirmed that thylakoid lipids (TL) were able to allosterically enhance modulator-induced LPOR activation. However, the allosteric modulation mechanism of LPOR by these compounds remains unclear. Herein, we integrated multiple computational approaches to explore the potential cavities in the Arabidopsis thaliana LPOR and an allosteric site around the helix-G region where high affinity for phosphatidyl glycerol (PG) was identified. Adopting accelerated molecular dynamics simulation for different LPOR states, we rigorously analyzed binary LPOR/PG and ternary LPOR/NADPH/PG complexes in terms of their dynamics, energetics, and attainable allosteric regulation. Our findings clarify the experimental observation of increased NADPH binding affinity for LPOR with PGs. Moreover, the simulations indicated that allosteric regulators targeting LPOR favor a mechanism involving lid opening upon binding to an allosteric hinge pocket mechanism. This understanding paves the way for designing novel LPOR activators and expanding the applications of LPOR. Full article
(This article belongs to the Special Issue Modulation of Protein Structure and Function by Lipids)
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Graphical abstract

Graphical abstract
Full article ">Figure 1
<p>Kinetic mechanism of <span class="html-italic">At</span>LPOR. Note ordered binding of Pchlide substrate and NADPH cofactor.</p> Full article ">Figure 2
<p>Structural formulas of phosphatidylglycerol (PG) and sulfoquinovosyldiacyglycerol (SQDG). The R groups and R’ represent various fatty acid chains of different lengths. <sup>a</sup> Stereospecifically numbering (Sn).</p> Full article ">Figure 3
<p>(<b>A</b>) Conformational superposition of <span class="html-italic">Arabidopsis thaliana</span> LPOR (PDB ID: 7JK9) and the constructed <span class="html-italic">At</span>LPOR in the “closed” (light green) and “open” (light yellow) state. The lid regions where the two proteins differ most are highlighted in green and orange, respectively. (<b>B</b>) The conformation of constructed <span class="html-italic">At</span>LPOR α-helices and β-sheets are shown in green and yellow, and connecting loop regions are shown in silver. (<b>C</b>) The constructed <span class="html-italic">At</span>LPOR/NADPH/Pchlide ternary complex.</p> Full article ">Figure 4
<p>(<b>A</b>) The ternary complexes obtained by molecular docking with PGs in three potential allosteric pockets. (<b>B</b>) The distance between PG and sites I, II, and III in three unbiased molecular dynamics simulations. (<b>C</b>) The most-hit site II structure superposition. The purple ball represents the site identified by PARS, the cyan surface represents the site identified by Corrsite, and the yellow spheres represent the site identified by MOLCAD. (<b>D</b>) Amplification of site II with the key residues around. For clarity, the solvent and hydrogen atoms are not shown. (<b>E</b>) The docking scores for PGs and SQDGs to the open and closed LPOR conformations. The curves defined by dispersed coordinate points for the total score are approximated by second-order polynomial fitting. (<b>F</b>) The docking mode for PGs and SQDGs in LPOR in the open state. PGs and SQDGs are shown as cyan and green stick models, respectively, and LPOR is shown with silver cartoons. The dashed yellow lines represent hydrogen bond interactions between lipids and the neighboring residues (shown as magenta lines). The enlarged image in the upper right corner highlights the hydrophilic interaction between multiple lipid heads with LPOR protein.</p> Full article ">Figure 5
<p>Electrostatic potential isosurface for (<b>A</b>) site II with the surrounding lid region, where red and blue represent positive and negative charges, respectively. (<b>B</b>) Schematic diagram of the electrostatic interaction driving PG to bind to site II. (<b>C</b>) PG (shown as cyan sticks) was captured to further promote the NADPH binding by affecting the lid conformation. (<b>D</b>) The total energy of LPOR binding PGs with different lengths. (<b>E</b>) Contribution of the energy term for favorable binding in nine systems (Ele: blue, Vdw: green, and Non-Pol: yellow, respectively). (<b>F</b>) The distribution of the electrostatic potential energy for different PG molecules on the electron density isosurfaces. The negatively charged nature of the PG head group caused the electrostatic potential energy of each part to be negative.</p> Full article ">Figure 6
<p>(<b>A</b>) Changes in the RMSD values of the lid relative to the RFD domain in the LPOR/PG structure during the simulations. (<b>B</b>) RMSF plots of all the atoms of LPOR/PG and apoLPOR. Structure data of lid1 (<b>C</b>) and lid2 (<b>D</b>) colored according to the opening pathway progress from yellow to green and orange to red, respectively. (<b>E</b>) Sampling of variables quantifying the lid position in aMD simulations of closed (red), open (blue) apoLPOR, and the open LPOR/PG (orange) systems. Dots represent MD snapshots in the last 20 ns runs. The variables are defined as two dihedral angles, representing the flipping of the I229-T230-G231 and 310I-311A-312T rotary microswitches in the hinge region. The details are shown in <a href="#app1-ijms-24-00307" class="html-app">Figure S16</a>. (<b>F</b>) The integral opening process shown to emphasize the hinge and lid motion in the simulated close LPOR structure, viewed from above. The black arrows display the movement direction of the lid in the simulations with respect to the inactive closed state. (<b>G</b>) Switching of the two salt bridges in the representative conformations of the LPOR/PG systems. The yellow dotted line represents the length of the salt bridge. (<b>H</b>) Comparison of the distances between the N atom in the side chain of K243/332 and O atom in the side chain of E379/343 for the LPOR/PG systems. (<b>I</b>) Distributions of distances 1 and 2 of Lys332-Glu343 and Lys243-Glu379 during the 50 ns simulation time for closed apoLPOR and LPOR/PG. The green, pink, blue, and yellow solid lines denote D1 and D2 in the closed crystal structure (PDB ID: 7JK9) and open structure of LPOR, respectively.</p> Full article ">Figure 7
<p>(<b>A</b>) The conformational transition extracted in LPOR/PG simulation. The PG is shown in stick with blue and yellow in MES and TES conformations, respectively. (<b>B</b>) The heat map of the decomposition of LPOR/PG in the binding pocket. The shades of red and blue represent the magnitude of BFE, respectively, and the energy items of electrostatic energy (ele), van der Waals (vdW), nonpolar, and polar contributions to solvation (sol) are shown in different color. (<b>C</b>) The RMSD curves for two binding orders. The NADPH (left) or PG (right) preferentially combined with LPOR, respectively. (<b>D</b>) Superimposition of the binding mode represented by PG binding mode cluster in LPOR/NADPH/PG (left) and in LPOR/PG/NADPH (right) with different conformations. Below each binding mode illustration, the ranking number of the binding mode corresponding to (<a href="#app1-ijms-24-00307" class="html-app">Table S11 and S12</a>) are listed. The TES conformation is obtained by manual alignment as a reference.</p> Full article ">
11 pages, 2095 KiB  
Article
Bioinformatic Analysis of Na+, K+-ATPase Regulation through Phosphorylation of the Alpha-Subunit N-Terminus
by Emma-Lucille Blayney, Milna Chennath, Charles G. Cranfield and Ronald J. Clarke
Int. J. Mol. Sci. 2023, 24(1), 67; https://doi.org/10.3390/ijms24010067 - 21 Dec 2022
Cited by 5 | Viewed by 1837
Abstract
The Na+, K+-ATPase is an integral membrane protein which uses the energy of ATP hydrolysis to pump Na+ and K+ ions across the plasma membrane of all animal cells. It plays crucial roles in numerous physiological processes, [...] Read more.
The Na+, K+-ATPase is an integral membrane protein which uses the energy of ATP hydrolysis to pump Na+ and K+ ions across the plasma membrane of all animal cells. It plays crucial roles in numerous physiological processes, such as cell volume regulation, nutrient reabsorption in the kidneys, nerve impulse transmission, and muscle contraction. Recent data suggest that it is regulated via an electrostatic switch mechanism involving the interaction of its lysine-rich N-terminus with the cytoplasmic surface of its surrounding lipid membrane, which can be modulated through the regulatory phosphorylation of the conserved serine and tyrosine residues on the protein’s N-terminal tail. Prior data indicate that the kinases responsible for phosphorylation belong to the protein kinase C (PKC) and Src kinase families. To provide indications of which particular enzyme of these families might be responsible, we analysed them for evidence of coevolution via the mirror tree method, utilising coevolution as a marker for a functional interaction. The results obtained showed that the most likely kinase isoforms to interact with the Na+, K+-ATPase were the θ and η isoforms of PKC and the Src kinase itself. These theoretical results will guide the direction of future experimental studies. Full article
(This article belongs to the Special Issue Modulation of Protein Structure and Function by Lipids)
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Figure 1

Figure 1
<p>Proposed operation of the electrostatic switch mechanism in the regulation of the Na<sup>+</sup>, K<sup>+</sup>-ATPase. The E2 conformation of the enzyme is stabilised through an electrostatic interaction between positively charged lysine residues on the N-terminus and the negatively charged headgroups of phosphatidylserine on the cytoplasmic surface of the surrounding membrane. Phosphorylation of hydroxyl groups of conserved serine and tyrosine residues of the N-terminus by protein kinases introduces negative charges onto the N-terminus, weakening its electrostatic interaction with the membrane and causing its release from the membrane. This destabilises the E2 conformation of the protein and facilitates its conformational change into the E1 state. The E2 and E1 structures shown in the figure were derived from published crystal structures PDB 3B8E in the case of the E2 state [<a href="#B7-ijms-24-00067" class="html-bibr">7</a>] and PDB 3WGU in the case of the E1 state [<a href="#B8-ijms-24-00067" class="html-bibr">8</a>].</p> Full article ">Figure 2
<p>Correlations of evolutionary distances between common organisms expressing various isoforms of PKC and the Na<sup>+</sup>, K<sup>+</sup>-ATPase α<sub>1</sub>-subunit. Each data point represents a combination of a pair of common organisms for each protein as explained in the Methods section. Hence, there are many more data points than common organisms. For comparison, the top row of the figure shows correlations of the positive control, PI3K-I<sub>A</sub>, with the Na<sup>+</sup>, K<sup>+</sup>-ATPase followed by two negative controls, i.e., PKCα, with EGFR followed by hexokinase with the Na<sup>+</sup>, K<sup>+</sup>-ATPase.</p> Full article ">Figure 3
<p>Correlations of evolutionary distances between common organisms that express different members of the Src kinase family of enzymes and the Na<sup>+</sup>, K<sup>+</sup>-ATPase α<sub>1</sub>-subunit.</p> Full article ">Figure 4
<p>Phylogenetic distributions of the Na<sup>+</sup>, K<sup>+</sup>-ATPase α<sub>1</sub>-subunit and PKCθ.</p> Full article ">
23 pages, 6739 KiB  
Article
Use of a Molecular Switch Probe to Activate or Inhibit GIRK1 Heteromers In Silico Reveals a Novel Gating Mechanism
by Dimitrios Gazgalis, Lucas Cantwell, Yu Xu, Ganesh A. Thakur, Meng Cui, Frank Guarnieri and Diomedes E. Logothetis
Int. J. Mol. Sci. 2022, 23(18), 10820; https://doi.org/10.3390/ijms231810820 - 16 Sep 2022
Cited by 4 | Viewed by 1685
Abstract
G protein-gated inwardly rectifying K+ (GIRK) channels form highly active heterotetramers in the body, such as in neurons (GIRK1/GIRK2 or GIRK1/2) and heart (GIRK1/GIRK4 or GIRK1/4). Based on three-dimensional atomic resolution structures for GIRK2 homotetramers, we built heterotetrameric GIRK1/2 and GIRK1/4 models [...] Read more.
G protein-gated inwardly rectifying K+ (GIRK) channels form highly active heterotetramers in the body, such as in neurons (GIRK1/GIRK2 or GIRK1/2) and heart (GIRK1/GIRK4 or GIRK1/4). Based on three-dimensional atomic resolution structures for GIRK2 homotetramers, we built heterotetrameric GIRK1/2 and GIRK1/4 models in a lipid bilayer environment. By employing a urea-based activator ML297 and its molecular switch, the inhibitor GAT1587, we captured channel gating transitions and K+ ion permeation in sub-microsecond molecular dynamics (MD) simulations. This allowed us to monitor the dynamics of the two channel gates (one transmembrane and one cytosolic) as well as their control by the required phosphatidylinositol bis 4-5-phosphate (PIP2). By comparing differences in the two trajectories, we identify three hydrophobic residues in the transmembrane domain 1 (TM1) of GIRK1, namely, F87, Y91, and W95, which form a hydrophobic wire induced by ML297 and de-induced by GAT1587 to orchestrate channel gating. This includes bending of the TM2 and alignment of a dipole of two acidic GIRK1 residues (E141 and D173) in the permeation pathway to facilitate K+ ion conduction. Moreover, the TM movements drive the movement of the Slide Helix relative to TM1 to adjust interactions of the CD-loop that controls the gating of the cytosolic gate. The simulations reveal that a key basic residue that coordinates PIP2 to stabilize the pre-open and open states of the transmembrane gate flips in the inhibited state to form a direct salt-bridge interaction with the cytosolic gate and destabilize its open state. Full article
(This article belongs to the Special Issue Modulation of Protein Structure and Function by Lipids)
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Figure 1

Figure 1
<p>Pharmacological actions of a molecular switch moiety are reproduced in sub-microsecond MD simulations in GIRK1/2. (<b>A</b>) The structures of GIRK1 specific modulators ML297, GAT1587, and the site of molecular switching region (off the pyrazole ring) that controls compound activity. (<b>B</b>) Equilibrium binding site for ML297 and GAT1587 following a 300 ns stochastic dynamics simulation in complex with PIP<sub>2</sub> and GIRK1/2. (<b>C</b>) Ion permeation pathway for a single ion taken from the ML297-PIP2-GIRK1/2 simulation. (<b>D</b>–<b>F</b>) Ions conducted, minimum gate distances, and normalized salt-bridge formation over the last 150 ns of the simulations are shown.</p> Full article ">Figure 2
<p>GAT1587 binding site along the TM helices in the GIRK1/2 heterotetramer.(<b>A</b>) Equilibrium binding pose for ML297 when complexed with PIP<sub>2</sub> GIRK1/2 after 300 ns of stochastic dynamics. (<b>B</b>) A 2D schematic representation of the compound’s binding pose depicting protein ligand interactions. (<b>C</b>) Equilibrium binding pose for GAT1587 when complexed with PIP<sub>2</sub> GIRK1/2 after 300 ns of stochastic dynamics. (<b>D</b>) A 2D schematic representation of the compounds binding pose depicting protein ligand interactions.</p> Full article ">Figure 3
<p>ML297 decreases and GAT1587 increases the angle between the outer and inner TM2 segments on each side of the flexible GIRK1-G169. (<b>A</b>) Comparison between TM2 of the ML297 (dark orange) and the GAT1587 (light orange) complexed with the GIRK1/2 systems. (<b>B</b>) The two hinge points that allow for TM2 bending. (<b>C</b>) Ramachandran plots for residues within TM2. I167 is highlighted in both. (<b>D</b>) Schematic representation of the movements of TM2 and a quantification of the bending around GIRK1-G169. (<b>E</b>) Expanded model including the nearby TM1 and a quantification of these effects. (<b>F</b>) Relative movements of the two transmembrane helices.</p> Full article ">Figure 4
<p>A hydrophobic wire (W95-Y91-F87) couples TM1 to TM2 at the level of the HBC gate (M180) and is stabilized by ML297 but not by GAT1587 to regulate the E141/D173-dependent conduction. (<b>A</b>) Protein structure with the pi-stack along TM1 highlighted. (<b>B</b>) Interaction distributions in the presence of ML297 or GAT1587. Pink denotes pi–pi interactions with an upper cutoff of 4 angstroms. Light pink denotes Van der Waals interactions with an upper cutoff of 3 angstroms. (<b>C</b>) Protein structure with the stabilizing hydrophobic residues of the GIRK2 subunit that can interact with the TM1 hydrophobic wire residues. (<b>D</b>) Interaction distance distributions of hydrophobic residues involved when either ML297 or GAT1587 is present. Light pink denotes Van der Waals interactions with an upper cutoff of 3 angstroms. (<b>E</b>) Protein structure (<b>E1</b>) with acidic residues that interact with a potassium ion (highlighted interactions) (<b>E2</b>). Dotted lines indicate a broken interaction between residues. The solid lines indicate the formation of an interaction. (<b>F</b>) Interaction distance distributions of TM1 hydrophobic chain residues with the two residues of the dipole between D141 and D173 of GIRK1 and S181 (next to G180) of GIRK2 in the presence of ML297 or GAT1587. Light green denotes dipole integrations with an upper cutoff of 4 angstroms. (<b>G</b>) Schematic representation of the charge relay network between W95 and Y91 of the TM1 hydrophobic chain with the two acidic residues.</p> Full article ">Figure 4 Cont.
<p>A hydrophobic wire (W95-Y91-F87) couples TM1 to TM2 at the level of the HBC gate (M180) and is stabilized by ML297 but not by GAT1587 to regulate the E141/D173-dependent conduction. (<b>A</b>) Protein structure with the pi-stack along TM1 highlighted. (<b>B</b>) Interaction distributions in the presence of ML297 or GAT1587. Pink denotes pi–pi interactions with an upper cutoff of 4 angstroms. Light pink denotes Van der Waals interactions with an upper cutoff of 3 angstroms. (<b>C</b>) Protein structure with the stabilizing hydrophobic residues of the GIRK2 subunit that can interact with the TM1 hydrophobic wire residues. (<b>D</b>) Interaction distance distributions of hydrophobic residues involved when either ML297 or GAT1587 is present. Light pink denotes Van der Waals interactions with an upper cutoff of 3 angstroms. (<b>E</b>) Protein structure (<b>E1</b>) with acidic residues that interact with a potassium ion (highlighted interactions) (<b>E2</b>). Dotted lines indicate a broken interaction between residues. The solid lines indicate the formation of an interaction. (<b>F</b>) Interaction distance distributions of TM1 hydrophobic chain residues with the two residues of the dipole between D141 and D173 of GIRK1 and S181 (next to G180) of GIRK2 in the presence of ML297 or GAT1587. Light green denotes dipole integrations with an upper cutoff of 4 angstroms. (<b>G</b>) Schematic representation of the charge relay network between W95 and Y91 of the TM1 hydrophobic chain with the two acidic residues.</p> Full article ">Figure 5
<p>The ML297-induced SH movement, as a result of the TM1 movement, drives changes in the CD loop interactions causing stabilization of the G-loop in the open conformation through its residue GIRK2-E315.2 liberating K188.1 that coordinates PIP<sub>2</sub> to stabilize the HBC gate in the open conformation. (<b>A</b>) A model of how TM1 modulates the slide helix (<b>A1</b>) and a quantification of these effects (<b>A2</b>). (<b>B</b>) Relative movements of the activated and inhibited SH regions. (<b>C</b>) Protein structure (<b>C1</b>) with residues that link the SH to the CD loop (<b>C2</b>). (<b>D</b>) Distance distributions of key residues that drive the channel into the active state. Dark yellow denotes charge–charge interactions with an upper cutoff of 4.0 angstroms. Light green denotes dipole charge interactions with an upper cut off of 4.0 angstroms. (<b>E</b>) Schematic outline of the changes in key residue interactions. (<b>F</b>) Protein structure (<b>F1</b>) with residues that link the SH to the CD loop (<b>F2</b>). (<b>G</b>) Distance distributions of residues that drive the channel into the active state. Dark yellow denotes charge–charge interactions with an upper cutoff of 4.0 angstroms. Pink denotes dipole charge interactions with an upper cutoff of 4.0 angstroms.</p> Full article ">Figure 5 Cont.
<p>The ML297-induced SH movement, as a result of the TM1 movement, drives changes in the CD loop interactions causing stabilization of the G-loop in the open conformation through its residue GIRK2-E315.2 liberating K188.1 that coordinates PIP<sub>2</sub> to stabilize the HBC gate in the open conformation. (<b>A</b>) A model of how TM1 modulates the slide helix (<b>A1</b>) and a quantification of these effects (<b>A2</b>). (<b>B</b>) Relative movements of the activated and inhibited SH regions. (<b>C</b>) Protein structure (<b>C1</b>) with residues that link the SH to the CD loop (<b>C2</b>). (<b>D</b>) Distance distributions of key residues that drive the channel into the active state. Dark yellow denotes charge–charge interactions with an upper cutoff of 4.0 angstroms. Light green denotes dipole charge interactions with an upper cut off of 4.0 angstroms. (<b>E</b>) Schematic outline of the changes in key residue interactions. (<b>F</b>) Protein structure (<b>F1</b>) with residues that link the SH to the CD loop (<b>F2</b>). (<b>G</b>) Distance distributions of residues that drive the channel into the active state. Dark yellow denotes charge–charge interactions with an upper cutoff of 4.0 angstroms. Pink denotes dipole charge interactions with an upper cutoff of 4.0 angstroms.</p> Full article ">Figure 6
<p>Overview of residue interactions driving the pre-open Apo channel state to the ML297-induced open conformation versus the GAT1587-induced close conformation. (<b>A</b>) A summary of the interaction networks that control channel function located near the compound binding site. (<b>B</b>) Outline of key changes affecting G loop open-state stabilization or a key Lys that coordinates PIP<sub>2</sub> to open the HBC gate.</p> Full article ">

Review

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17 pages, 2700 KiB  
Review
Molecular Dynamics Simulations of Curved Lipid Membranes
by Andreas Haahr Larsen
Int. J. Mol. Sci. 2022, 23(15), 8098; https://doi.org/10.3390/ijms23158098 - 22 Jul 2022
Cited by 14 | Viewed by 4074
Abstract
Eukaryotic cells contain membranes with various curvatures, from the near-plane plasma membrane to the highly curved membranes of organelles, vesicles, and membrane protrusions. These curvatures are generated and sustained by curvature-inducing proteins, peptides, and lipids, and describing these mechanisms is an important scientific [...] Read more.
Eukaryotic cells contain membranes with various curvatures, from the near-plane plasma membrane to the highly curved membranes of organelles, vesicles, and membrane protrusions. These curvatures are generated and sustained by curvature-inducing proteins, peptides, and lipids, and describing these mechanisms is an important scientific challenge. In addition to that, some molecules can sense membrane curvature and thereby be trafficked to specific locations. The description of curvature sensing is another fundamental challenge. Curved lipid membranes and their interplay with membrane-associated proteins can be investigated with molecular dynamics (MD) simulations. Various methods for simulating curved membranes with MD are discussed here, including tools for setting up simulation of vesicles and methods for sustaining membrane curvature. The latter are divided into methods that exploit scaffolding virtual beads, methods that use curvature-inducing molecules, and methods applying virtual forces. The variety of simulation tools allow researcher to closely match the conditions of experimental studies of membrane curvatures. Full article
(This article belongs to the Special Issue Modulation of Protein Structure and Function by Lipids)
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Figure 1

Figure 1
<p>Eukaryotic cell and organelles, with examples of various membrane curvatures. Some curvature-inducing proteins are also shown, see the main text.</p> Full article ">Figure 2
<p>Simulating membrane curvature induction. (<b>A</b>) Bending of plane periodic bilayer by protein crowding. Figure adapted with permission from Ref. [<a href="#B24-ijms-23-08098" class="html-bibr">24</a>] (copyright 2019, PNAS license). (<b>B</b>) Bicelle to vesicle transition, adapted from Ref. [<a href="#B25-ijms-23-08098" class="html-bibr">25</a>]. (<b>C</b>) Tethering from a plane membrane, adapted from Ref. [<a href="#B26-ijms-23-08098" class="html-bibr">26</a>]. If not stated otherwise, the original figures are published under a creative common license.</p> Full article ">Figure 3
<p>Methods for vesicle generation. (<b>A</b>) Self-assembly of vesicle, adapted from Ref. [<a href="#B35-ijms-23-08098" class="html-bibr">35</a>]. (<b>B</b>) CHARMM-GUI Martini Maker, adapted from Ref. [<a href="#B36-ijms-23-08098" class="html-bibr">36</a>]. Top shows the cross-section of a vesicle with pores for lipid translocation between leaflets, bottom shows the vesicle after pore-closure. (<b>C</b>) Generation of vesicles by triangular surfaces, using TS2CG. Adapted with permission from Ref. [<a href="#B5-ijms-23-08098" class="html-bibr">5</a>] (copyright 2015, ACS Publications). (<b>D</b>) Curvature generated from transition of a flat membrane, using BUMPy. Adapted from Ref. [<a href="#B37-ijms-23-08098" class="html-bibr">37</a>]. (<b>E</b>) Insertion of membrane protein in a vesicle, using TS2CG. Adapted from Ref. [<a href="#B5-ijms-23-08098" class="html-bibr">5</a>]. If not stated otherwise, the original figures are published under a creative common license.</p> Full article ">Figure 4
<p>Methods for setting up and maintaining curvature. (<b>A</b>) Scaffolding approach, employing a ‘wall’ of virtual beads (orange) that sustain membrane curvature by repulsive forces between the beads and lipid tails. Pure POPC membrane, set-up using TS2CGv1.1 [<a href="#B43-ijms-23-08098" class="html-bibr">43</a>]. The semi-transparent membrane is the periodic extension of the simulated box. (<b>B</b>) Scaffolding approach, where the membrane is attracted to a solid support with a hole. A cross-section through the hole is shown. Membrane of DOPC (blue) and POPI<sub>P2</sub> (red). Set-up using the protocol in Ref. [<a href="#B50-ijms-23-08098" class="html-bibr">50</a>]. (<b>C</b>) Lipid asymmetry-induced curvature. The anisotropy is maintained by virtual repulsive forces between POPC (blue) and POPS (red). Membrane fluctuations are suppressed by harmonic position restraints. Set up following Ref. [<a href="#B54-ijms-23-08098" class="html-bibr">54</a>]. (<b>D</b>) Virtual forces can generate and maintain a membrane curvature defined by a set of collective variables. Here, the virtual forces are harmonic restraints on the distance between a virtual bead (orange) and a section (yellow) of a POPC membrane. Set-up using EnCurv [<a href="#B48-ijms-23-08098" class="html-bibr">48</a>]. (<b>E</b>) The membrane is compressed in one direction (buckling) and then the box dimensions are fixed. Membrane fluctuations are avoided by position restraints. Mixed membrane of POPC (blue), POPS (red) and cholesterol (brown). Set up using Refs. [<a href="#B55-ijms-23-08098" class="html-bibr">55</a>,<a href="#B56-ijms-23-08098" class="html-bibr">56</a>]. (All original figures, generated for this review).</p> Full article ">Figure 5
<p>Experiments for measuring membrane curvature. (<b>A</b>) Electron microscopy images of how amphyphysin (wild type and mutant “mut2”) induce transitions from vesicles towards tubules. Adapted with permission from Ref. [<a href="#B75-ijms-23-08098" class="html-bibr">75</a>] (copyright 2004, The American Association for the Advancement of Science). (<b>B</b>) Fluorescent liposomes (red, middle panel) on a surface with bound BAR domains (green, right panel). Adapted with permission from Ref. [<a href="#B78-ijms-23-08098" class="html-bibr">78</a>] (copyright 2009, John Wiley and Sons). (<b>C</b>) Tethering from GUV with optical tweezer, adapted with permission from Ref. [<a href="#B79-ijms-23-08098" class="html-bibr">79</a>] (copyright 2014, Elsevier). (<b>D</b>) Simulated tethering and fission event, adapted with permission from Ref. [<a href="#B60-ijms-23-08098" class="html-bibr">60</a>] (copyright 2014, The American Association for the Advancement of Science).</p> Full article ">
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